RESEARCH ARTICLE

Molecular Reproduction & Development 79:296–307 (2012)

The Effect of Leptin on Maturing Porcine OocytesIs Dependent on Glucose Concentration

ELENA SILVA, MELISSA PACZKOWSKI,† AND REBECCA L. KRISHER†*

Department of Animal Sciences, University of Illinois at Urbana-Champaign, Urbana, Illinois

SUMMARY

Increased body weight is often accompanied by increased circulating levels of leptinand glucose, which alters glucose metabolism in various tissues, including perhapsthe oocyte. Alteration of glucose metabolism impacts oocyte function and maycontribute to the subfertility often associated with obese individuals. The objectiveof this study was to determine the effect of leptin (0, 10, and 100 ng/ml) on the oocyteand cumulus cells during in vitro maturation under differing glucose concentrations.Weexamined the effects of leptin on oocytematuration, blastocyst development, and/or gene expression in oocytes and cumulus cells (IRS1, IGF1,PPARg , IL6,GLUT1) ina physiological glucose (2mM) and high glucose (50mM) environment. We alsoevaluated the effect of leptin on glucose metabolism via glycolysis and the pentosephosphate pathway. In a physiological glucose environment, leptin did not have aninfluence on oocyte maturation, blastocyst development, or oocyte gene expression.Expression of GLUT1 in cumulus cells was downregulated with 100 ng/ml leptintreatment, but did not affect oocyte glucose metabolism. In a high glucose environ-ment, oocyte maturation and glycolysis were decreased, but in the presence of100 ng/ml leptin, these parameters were improved to levels similar to control. Thiseffect is potentially mediated by an upregulation of oocyte IRS1 and a correction ofcumulus cell IGF1 expression. The present study demonstrates that in a physiologicalglucoseconcentration, leptin plays anegligible role in oocyte function.However, leptinappears to modulate the deleterious impact of a high glucose environment on oocytefunction.

Mol. Reprod. Dev. 79: 296–307, 2012. � 2012 Wiley Periodicals, Inc.

Received 29 December 2011; Accepted 27 January 2012

* Corresponding author:National Foundation for FertilityResearch

10290 RidgeGate CircleLone Tree, CO 80124.E-mail: rkrisher@fertilityresearch.org

† Current address:National Foundation for FertilityResearch

Lone Tree, CO 80124.

Grant sponsor: University of Illinois,College of Agriculture, Consumer andEnvironmental Sciences

Published online 14 February 2012 in Wiley Online Library(wileyonlinelibrary.com).DOI 10.1002/mrd.22029

INTRODUCTION

Increased body weight is associated with anovulation,subfertility, and increased risk of pregnancy loss (Brewerand Balen, 2010). The adipokine leptin is produced mainlyby the adipocyte, and its primary function is to controlfood intake and energy metabolism (Halaas et al., 1995;Pelleymounter et al., 1995). Leptin is also responsible forcommunication between adipose tissue and the reproduc-tive system. Leptin-deficientmice are infertile (Barash et al.,1996), demonstrating the important interaction betweenbody mass and reproductive function mediated by leptin.

Both in vitro and in vivo studies demonstrate that leptinregulates the hypothalamic-pituitary-ovarian axis viastimulation of gonadotropin-releasing hormone, follicle-stimulating hormone, and luteinizing hormone release(Yu et al., 1997; Watanobe, 2002).

Abbreviations: #G#L, glucose (mM), leptin (ng/ml) concentrations; COCs,cumulus-enclosed oocyte complexes; GLUT1, glucose transporter 1; IGF1,insulin-like growth factor 1; IL6, interleukin-6; IRS1, insulin receptor substrate1; IVM, in vitro maturation; PPARg, peroxisome proliferator-activated receptorgamma; PPP, pentose phosphate pathway.

� 2012 WILEY PERIODICALS, INC.

Serum and follicular fluid leptin levels have a positivecorrelation with body mass index; obese individuals havegreater leptin concentrations than lean individuals(Considine et al., 1996). Leptin has been suggested asan indicator of fertility, with serum concentrations greaterthan 59 ng/ml associated with reduced embryo quality(Anifandis et al., 2005). The link between obesity andinfertility has been associated with hyperleptinemia andalterations of the hypothalamic-ovarian axis (Tortorielloet al., 2004), resulting in impairment of ovulation (Duggalet al., 2000) and reduced follicular growth (Swain et al.,2004). Studies regarding leptin’s direct effect on theoocyte are contradictory, however, with either a positiveor no effect of leptin on nuclear maturation and subsequentblastocyst development described in pig (Craig et al.,2004; Jin et al., 2009), mouse (Ryan et al., 2002; Swainet al., 2004; Ye et al., 2009), and bovine (Cordova et al.,2011) oocytes. In addition to a direct effect on the oocyte,leptin’s influence on oocyte maturation may be alsocumulus cell-mediated (Paula-Lopes et al., 2007; vanTol et al., 2008). Because leptin and leptin receptormRNA are present in both the oocyte and cumulus cells(Cioffi et al., 1997; Ryan et al., 2002; Craig et al., 2004;Benomar et al., 2006; Arias-Alvarez et al., 2010), leptinmayplay an important role in regulating oocyte function andcompetence.

In addition to an increase in leptin in obese individuals,serum glucose is also elevated, as obesity is often asso-ciated with altered insulin signaling in nonreproductivetissues (Matthaei et al., 2000). Elevation of glucose levelsin obese women extends to the ovary, where there is anincrease in follicular fluid glucose concentration (Robkeret al., 2009). Exposure of oocytes to elevated glucose iscorrelated with delayed nuclear maturation and reducedblastocyst development (Diamond et al., 1989; Hashimotoet al., 2000; Wang et al., 2009). The deleterious effect ofhigh glucose on oocyte competence could be mediatedby inhibition of glucose metabolism, as reduced glycolysisis known to decrease oocyte developmental competence(Krisher and Bavister, 1999; Spindler et al., 2000; Herricket al., 2006). Increased follicular glucose may alsoinduce a downregulation of oocyte glucose transporters,as is the case in mouse embryos (Moley et al., 1998b). Asystemic increase in glucose concentrations, as in dia-betes, impairs the oocyte directly and also alters the inter-action between the oocyte and its surrounding cumuluscell mass (Colton et al., 2003), which also likely impactsoocyte quality.

Oocytes metabolize glucose via multiple pathwaysincluding glycolysis, the pentose phosphate pathway(PPP), and the Krebs cycle (Rieger and Loskutoff,1994; Downs et al., 1998; Krisher and Bavister, 1999). Itis important to note that there are species differencesregarding glucose preferences during in vitro oocytematuration. In pigs, glucose is required for optimal oocytematuration and developmental competence (Funahashiet al., 2008; Silva and Krisher, 2008), whereas in mice,glucose is not necessary to sustain oocyte maturation ifmedium is supplemented with pyruvate (Downs and

Mastropolo, 1994). This difference highlights the impor-tance of glucose for the pig oocyte, as well as its preferencefor glycolysis and PPP over the Krebs cycle (Herrick et al.,2006; Krisher et al., 2007). Despite species differences inglucose requirements, an increase in glucose metabolismvia the different metabolic pathways occurs during meioticresumption inmice (Downset al., 1996), bovine (Rieger andLoskutoff, 1994), and cat (Spindler et al., 2000). Yet, toomuch available glucose is deleterious to the oocyte, due toimpairment of oocyte metabolism (Ratchford et al., 2007)and increased oxidative stress (Hashimoto et al., 2000),emphasizing the importance of adequate glucose levels foroptimal oocyte quality.

Given that circulating levels of leptin and glucose areinfluenced by body weight, it is important to determineleptin’s role in oocyte function in differing glucose concen-trations. The interaction between leptin and glucose hasbeen extensively studied in peripheral tissues. Leptinand insulin signaling pathways crosstalk to regulateglucosemetabolism (Koch et al., 2010) and glucose uptakeand transport in neuronal, hepatic, and muscle cells(Yaspelkis et al., 2002; Lam et al., 2004; Benomar et al.,2006). Yet, chronic exposure to high glucose and leptininduces apoptosis and impairs glucose-stimulated insulinsecretion in pancreaticb cells (Maedler et al., 2008). Limitedinformation is available about how leptin and glucosemeta-bolism interact to affect oocyte and cumulus cell function.The objective of this study was to determine the effect ofleptin on oocytes and cumulus cells during in vitro matura-tion (IVM) with variable glucose concentrations. Becauseleptin signaling is altered in conditions of obesity andhyperglycemia (Tortoriello et al., 2004), we hypothesizethat the effect of leptin on oocyte maturation and metabo-lism differs in physiological and supraphysiological glucoseenvironments.

RESULTS

Effect of Leptin on Oocytes and Cumulus CellsMatured in a Physiological Glucose Environment

Leptin supplementation (0, 10, and 100ng/ml) duringoocyte maturation in a physiological glucose (2mM, 2G)environment did not affect the percentage of oocytessuccessfully completing meiotic maturation (P¼ 0.16,Table 1). Additionally, leptin supplementation during oocytematuration had no effect on subsequent embryonic clea-vage (P¼0.09), blastocyst development (P¼ 0.51), or totalblastocyst cell number (P¼ 0.57, Table 1). Transcript abun-dance of insulin receptor substrate 1 (IRS1), insulin-likegrowth factor 1 (IGF1), peroxisome proliferator-activatedreceptor gamma (PPARg), and interleukin-6 (IL6) werenot significantly different in oocytes or cumulus cellsmatured with or without leptin at any concentration(Fig. 1). The expression of glucose transporter 1(GLUT1) was also analyzed in cumulus cells; GLUT1was downregulated with the addition of 100 ng/ml leptinduring maturation compared to the control treatment with-out leptin (Fig. 1).

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Effect of Leptin on Oocytes and Cumulus CellsMatured in a Supraphysiological GlucoseEnvironment

In the absence of leptin, control treatment (standardglucose for maturation media, 5mM, 5G0L) resulted in ahigher percentage of successful nuclear maturation com-pared to a supraphysiological glucose (50mM, 50G0L)environment (83.4% vs. 71.7%, respectively; P¼0.03,Table 2). Interestingly, leptin supplementation (100ng/ml)during in vitromaturation (IVM) of porcine cumulus-enclosedoocyte complexes (COCs) in a supraphysiological glucoseenvironment significantly improved nuclear maturation com-

pared to 50G0L (80.4% vs. 71.7%, respectively; P¼0.04;Table 2), with leptin minimizing the deleterious effect of highglucose. Lower concentrations of leptin (10ng/ml, 50G10L)also improved nuclear maturation to levels similar to control;however, this treatment was not significantly different than50G0L (79.8% vs. 71.1%, P¼ 0.23).

We examined transcript abundance of the same genesanalyzed in the previous experiment in oocytes and cumu-lus cells. There was no detectable expression of IGF1 in theoocytes sampled in this experiment, possibly due to natu-rally occurring oocyte variability or the difference in numberof oocytes in each sample pool in this experiment (20oocytes) as compared to the previous experiment (30oocytes). Porcine oocyte expression of IGF1 decreasesduring oocyte maturation (Zhu et al., 2008). A previousstudy using a greater number of bovine oocytes also did notdetect expression of IGF1 (Warzych et al., 2007). A 10-foldincrease in glucose alone did not result in a significantdifference in oocyte gene expression for any of the genesanalyzed (Fig. 2).Within the high glucose treatment groups,100 ng/ml leptin upregulated oocyte expression of IRS1. Incontrast, 10 ng/ml leptin significantly downregulated IL6expression compared to control treatment (5G0L), andsignificantly downregulated PPARg expression comparedto all other treatment groups (Fig. 2). In cumulus cells, a 10-fold increase in glucose alone upregulated the expressionof IGF1 in comparison to control treatment; expression ofthe other genes analyzed (IRS1, IL6, PPARg, and GLUT1)was not different. Within the high glucose treatmentgroups, addition of leptin during COC maturation had theopposite effect overall onexpressionof thegenesanalyzed.Treatment with 10 ng/ml leptin downregulated expression

TABLE 1. Effect of Leptin Supplementation During Porcine Oocyte IVM in a Physiological Glucose (2mM) Environment onNuclear Maturation and Subsequent Embryonic Cleavage, Blastocyst Development, and Blastocyst Total Cell Number

Following In Vitro Fertilization*

Leptin (ng/ml) Maturation, % (n) Cleavage, % (n) Blastocyst/total, % (n) Blastocyst/cleaved (%) Blastocyst cell number (n)

0 79.4� 6.2 (155) 63.5� 5.8 (128) 20.1� 5.3 (128) 34.8� 11.7 51.2� 3.9 (21)10 73.3� 6.4 (145) 54.9� 9.0 (122) 16.7� 6.7 (122) 33.6� 15.2 55.3� 5.1 (18)100 82.0� 4.8 (153) 74.6� 8.1 (145) 23.4� 3.1 (145) 32.9� 6.6 49.2� 4.9 (20)

n¼Total number of oocytes/embryos in each group.

*Data are reported as mean�SEM. No significant differences were observed between the variables analyzed in each column.

Figure 1. Oocyte (a) and cumulus cell (b) relative expression of thetarget genes IRS1, IGF1, IL6, PPARg, and GLUT1 following leptinsupplementation during IVM in physiological glucose conditions(2mM, 2G), as determined by qPCR analysis. Bars represent genetranscript abundance in each treatment relative to the control treat-ment (2G0L). Data were normalized to the expression of each gene inthe control treatment, which was assigned a value of 1. Columns withdifferent superscript letterswithin eachgene differ (a,b;P�0.05).Nosignificant difference was detected between treatments in oocytes.

TABLE 2. Effect of Leptin Supplementation During IVM onthe Percentage ofOocytes CompletingNuclearMaturation in

a Supraphysiological Glucose Environment (50mM)

Treatment

Mature (%)1Glucose (mM) Leptin (n)

5 0 ng/ml (249) 83.4� 3.1a

50 0 ng/ml (213) 71.7� 5.1b

50 10 ng/ml (238) 79.8� 5.1ab

50 100ng/ml (206) 80.4� 4.6a

n¼Number of oocytes in each treatment group.1Values presented are mean�SEM of oocytes matured in 5–8 replicates.

Different superscript letters within a column differ (P< 0.05).

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of IRS1 and PPARg, while treatment with 100 ng/ml leptindownregulated expression of IRS1, IGF1, and IL6 in cumu-lus cells (Fig. 2). In contrast to the effect of leptin onGLUT1expression in cumulus cells in physiological glucose con-ditions, leptin treatment had no effect on cumulus cellGLUT1 expression in a high glucose maturationenvironment.

Effect of Leptin During In Vitro Maturation onOocyte Glucose Metabolism

Leptin supplementation (0, 10, and 100ng/ml) duringoocyte maturation in the standard medium glucose con-centration (5mM) had no effect on glycolysis or PPPactivity(Table 3). An increase in glucose concentration duringCOCmaturation from 5 to 50mM reduced glycolytic activity(P¼0.04; Table 4); however, within the 50mM glucosetreatments, there was no effect of leptin on glycolysis.Treatment with 100 ng/ml leptin restored glycolytic levelsequal to that in standard glucose (P>0.10), while treatmentwith 10 ng/ml leptin still tended to reduce glycolytic activitycompared to the standard glucose control (P¼0.06).Neither the increase in glucose alone nor leptin treatment

in a high glucose environment had any effect on PPPactivity (P¼0.30).

DISCUSSION

These results demonstrate that the porcine oocyte isresponsive to leptin, and that the response elicited dependson the glucose concentration in the maturation environ-ment. Leptin had a limited influence on oocyte function in aphysiological glucose concentration, whereas leptin mini-mized the deleterious effect of a high glucose environmentwith an improvement in both oocyte nuclear maturation andglucose metabolism, potentially mediated by an upregula-tion of oocyte IRS1 and a correction of cumulus cell IGF1expression.

The leptin receptor is expressed at all stages of porcineoocytes duringmaturation, although it is significantly higherat germinal vesicle breakdown, and binding influencesoocyte maturation via the mitogen-activated protein kinasepathway (Craig et al., 2004). While activation of theleptin pathway was not directly examined in our study,the alteration of gene expression, nuclear maturation,and glycolysis that we observed in a high glucose environ-ment indicates that leptin was active in our system. Ourfinding that leptin does not influence oocyte developmental

Figure 2. Oocyte (a) and cumulus cell (b) relative expression ofthe target genes IRS1, IGF1, IL6, PPARg, andGLUT1 following leptinsupplementation during IVM in supraphysiological glucose conditions(50mM; 50G), as determined by qPCR analysis. Oocyte expressionof IGF1was not detected in this experiment. No significant differencewas detected for GLUT1 expression in cumulus cells. Bars representgene transcript abundance in each treatment relative to the controltreatment (5G0L). Data were normalized to the expression ofeach gene in the control treatment, which was assigned a value of1. Columns with different superscript letters within each gene differ(a,b; P�0.05). *Tended to differ (P¼0.06).

TABLE 3. Effect of Leptin Supplementation During PorcineOocyte IVM in Standard-Glucose (5mM) Conditions onOocyte Glycolytic and Pentose Phosphate Pathway

(PPP) Activity*

Treatment

Glycolysis�SE(pmol/oocyte/3 hr)

PPP�SE(pmol/oocyte/3 hr)

Glucose(mM) Leptin (n)

5 0 ng/ml (47) 1.42� 0.10 0.44� 0.045 10 ng/ml (40) 1.32� 0.11 0.41� 0.045 100ng/ml (54) 1.29� 0.08 0.43� 0.03

n¼Number of oocytes in each treatment group.

*No significant differences were observed between the variables analyzed in

each column.

TABLE 4. Effect of Leptin Supplementation During PorcineOocyte IVM in Supraphysiological Glucose (50mM)

Conditions on Oocyte Glycolytic and PentosePhosphate Pathway (PPP) Activity

Treatment

Glycolysis�SE(pmol/oocyte/3 hr)

PPP�SE(pmol/oocyte/3 hr)

Glucose(mM) Leptin (n)

5 0 ng/ml (55) 1.49� 0.10a* 0.53� 0.0450 0 ng/ml (42) 1.08� 0.10b 0.44� 0.0650 10 ng/ml (34) 1.16� 0.11a,b * 0.48� 0.0950 100ng/ml (42) 1.27� 0.13a,b 0.49� 0.10

n¼Number of oocytes in each treatment group.a,bDifferent superscript letters within a column differ (P<0.05).

*Tended to differ (P¼0.06).

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competence duringmaturation under physiological glucoseconditions is in agreement with previous studies in pigoocytes (Jin et al., 2009; Suzuki et al., 2010). In contrast,Craig et al. (2004) observed a positive effect of leptin (at10 and 100 ng/ml) on nuclear maturation of pig oocytes.Maturationof the control group in our studywasgreater thanthat reported by Craig et al. (79.2% vs. 65.9%), so it ispossible that under optimal in vitro conditions, leptin treat-ment doesnot further improveoocytenuclearmaturation. Inaddition, ovary source (gilt vs. sow) andmaturationmedium(Tissue Culture Medium 199 vs. Purdue Porcine Mediummodified for maturation) differed between Craig et al. andour study, and possibly influenced the results obtained.Variation in animal metabolic status also influences leptinfunction (Barb et al., 2008); as nutritional requirementsdiffer between gilts and sows in a production setting, animalagemay influence the parameters examined. In agreementwith our maturation and development data, leptin treatmenthad no effect on oocyte expression of IRS1, IGF1, PPARg ,or IL6 in a physiological glucose environment. A high leptinconcentration (100 ng/ml) downregulated expression ofGLUT1 in cumulus cells when added during IVM in physio-logical glucose. There is well-established metabolic coop-erativity between the oocyte and cumulus cells, as thecumulus cells metabolize significant amounts of glucoseand transport lactate and pyruvate to the oocyte throughgap junctions (Su et al., 2009). Because an increase inGLUT1 expression is correlatedwith an increase in glucoseincorporation in mouse embryos (Morita et al., 1994), it isexpected that downregulation of GLUT1 expression wouldbe associated with a reduction in glucose uptake andalteration of glucose metabolism in the COC. This appearsto be a metabolic perturbation specifically in the cumuluscells as there was no effect of leptin on the glycolyticpathway or PPP when oocytes were matured in a similarlow glucose environment (5mM). Previous work did notdemonstrate a difference in glycolytic pathway activitybetween 2 and 5mM glucose (Krisher et al., 2007). Thissupports our conclusion that downregulation of GLUT1 inthe cumulus cells had no impact on oocyte glycolyticactivity.

As expected, a 10-fold increase in glucose concentra-tion, irrespective of leptin, had a negative impact on oocytenuclear maturation. This is in agreement with a previousreport in diabetic mice where hyperglycemia also causedimpairment of oocyte maturation (Colton et al., 2002). Invitro exposure of mouse embryos and oocytes to a similarhigh glucose concentration (52mM) mimics the in vivoeffect of diabetic conditions (Moley et al., 1998a; Adastraet al., 2011). In previouswork, an increase in glucose from2to 10mM also significantly reduced glycolytic and PPPactivity in mature porcine oocytes, possibly due to inhibitionof the enzymes present in the glycolytic pathway (Krisheret al., 2007). The tendency for downregulation of GAPDHexpression that we observed in oocytes exposed to 50mMglucose (data not shown) supports this hypothesis. Despitethe effect on oocyte meiotic maturation and glycolysis,supraphysiological glucose concentration had no impacton oocyte expression of the genes analyzed compared to

standard glucose. There were modest changes in cumuluscell gene expression, with upregulation of IGF1 expression.IGF1 is a known promoter of cumulus cell expansion (Singhand Armstrong, 1997); however, a supraphysiological con-centration of IGF1 protein has deleterious effects onembryos via downregulation of IGF1 receptor, causingreduced glucose uptake and apoptosis (Chi et al., 2000).We do not know if the altered IGF1 transcript level incumulus cells that we observed resulted in a change inIGF1 protein level; further studies are necessary to confirmthese results.

In contrast to the limited effect of leptin onoocyte functionin a physiological glucose condition, leptin had significanteffects on oocyte nuclear maturation, oocyte glucosemetabolism, and oocyte and cumulus cell gene expressionin a supraphysiological glucose environment. High leptinconcentration (100 ng/ml) significantly improved nuclearmaturation and glycolysis in oocytes exposed tosupraphysiological glucose to levels similar to the standardglucose control, demonstrating that leptin minimizes thenegative effects of high glucose. Our data suggest that thispositive effect is likely mediated by changes in oocyte andcumulus cell gene expression, and consequent improve-ment of glycolytic activity. The upregulation by leptin ofoocyte IRS1 expression may have a positive impact onglycolytic activity since IRS1 activation transmits signalfrom insulin and IGF1 receptors and triggers phosphatidy-linositol 3-kinase activation, thereby modulating glucosemetabolism in tissues including the ovary (Poretskyet al., 1999). Leptin directly regulates IRS1 in several othercell types (Cohen et al., 1996; Hennige et al., 2006; Yanget al., 2009). A high leptin concentration was necessary tosignificantly improve glycolysis; 10 ng/ml tended to havelower glycolytic activity than control, resulting in an inter-mediate effect. Ten ng/ml of leptin also failed to significantlyupregulate oocyte IRS1 expression and downregulated IL6and PPARg expression. IL6 has not been identified as aregulator of glucose metabolism in the ovary, but IL6enhances glucose transport in adipocytes and muscle cells(Stouthard et al., 1996; Glund et al., 2007). Also, IL6 andglucose metabolism are integral to cumulus cell expansionduring oocytematuration (Sutton-McDowall et al., 2006; Liuet al., 2009). Perturbations of oocyte–cumulus cell signalingaffect oocyte function, and because the cumulus cellsmetabolize a large percentage of the glucose available,alterations in the IL6 pathwaymay influence glucose uptakeby the cumulus cells as well as its metabolic fate. Themechanism of PPARg regulation of insulin sensitivity inthe ovary is still unknown; however, PPARg improvesglucose homeostasis in other cell types, and its functionhas been associated with leptin transcriptional regulation(De Vos et al., 1996; Kim and Ahn, 2004). The positiveeffect of 100 ng/ml leptin on the oocyte in a supraphysiolo-gical environment is possibly mediated via upregulation inoocyte IRS1expressionaswell as rescueof theoocyte fromthedownregulation of IL6 andPPARg observedwith a lowerleptin concentration.

In contrast to the situation in oocytes, 100 ng/ml leptindownregulated cumulus cell expression of IRS1, IGF1, and

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IL6 compared to high glucose without leptin in a supraphy-siological glucose environment. Downregulation of thesegenes in cumulus cells following leptin treatmentmay play arole in improving oocyte function. In the case of IGF1, leptincorrects the upregulation caused by the supraphysiologicalglucose environment alone. As discussed previously, anincrease in IGF1 may have a deleterious effect on glucosemetabolism (Chi et al., 2000). Interestingly, the downregu-lation observed in IGF1, IRS1, and IL6 gene expression inthe cumulus cells had little impact on oocyte glucose meta-bolism; although glycolysis increased with increasingleptin, these differences were not significant within thehigh glucose group. Since our study only determinedglucose metabolism in denuded oocytes, we cannotexclude the possibility that differences in glycolysis and/or PPP activity may have been detected if oocyte glucosemetabolism could be evaluated in intact COCs. In addition,the lower leptin concentration downregulated PPARgexpression in cumulus cells similar to that in the oocyte,supporting the hypothesis that an alteration in PPARgexpression may compromise glycolysis in a high glucoseenvironment.

Taken together, these results indicate a positive effect ofleptin on oocyte competence in a supraphysiological glu-cose environment. Yet, the specific mechanism of thismodulation of oocyte function by leptin remains unclear.Leptin reduces systemic glucose levels and regulates glu-cose utilization in a diabetic mouse model via an insulinindependent pathway (Chinookoswong et al., 1999; Hidakaet al., 2002; Kojima et al., 2009). Hyperleptinemia reverseshyperglycemia in insulin-deficient mice via reduction ofglucagon and increased insulin sensitivity (Denrocheet al., 2011). We know that leptin directly regulates ovariansteroidogenesis and folliculogenesis by actingon the insulinsignaling pathway in theca and granulosa cells (Zachowand Magoffin, 1997; Kendall et al., 2004; Munoz-Gutierrezet al., 2005), but the effect of leptin on oocyte insulinsignaling is unclear. Our data suggest that leptin amelio-rates the deleterious effects of high glucose, at least in part,via alterations in oocyte glycolysis. But we cannot saywhether the increased oocyte glycolysis that we observedis responsible for enhanced oocyte viability in hyperglyce-mic conditions or is simply an indirect indicator of an oocytemade more viable via another mechanism. Because therewere only moderate differences in gene expressionbetween standard and supraphysiological glucose environ-ments, factors other than the ones examined here mediat-ing the impairment of oocyte physiology by high levels ofglucose are probably involved. One potential player isapoptosis as hyperglycemia triggers apoptosis in mouseembryos and oocytes (Moley et al., 1998a; Chang et al.,2005). Leptin reduces bovine cumulus cell (Paula-Lopeset al., 2007) and blastocyst apoptosis (Boelhauve et al.,2005) via upregulation of the apoptosis-related genes FASand BIRC4. We have observed that supplementation withhigh (100 ng/ml), but not low (10 ng/ml), leptin also upre-gulates oocyte FAS expression in the pig (data not shown).Therefore, in a high glucose environment leptin may alle-viate apoptosis, thus rescuing oocyte quality.

In summary, leptin’s effect on oocytephysiology is depen-dent on glucose concentrations during maturation. Leptintreatment appears to ameliorate the negative effect of highglucose on oocyte quality. This is an important finding as itexpands our knowledge regarding leptin function in thefollicular environment. These results indicate that, similarto nonreproductive tissues, leptin and glucose pathwaysinteract in the ovarian environment. Although ovarian leptininsensitivity has not been reported, systemic leptin resis-tance has been associated with infertility (Tortoriello et al.,2004; Brannian et al., 2005). The protective effect of leptinis possibly compromised in vivo in a hyperglycemic/hyperleptinemic environment due to leptin resistance. Nor-malization of the leptin and glucose pathways in the follicularenvironment, including the oocyte, may result in potentialtargets for infertility treatment. Moreover, since leptin differ-ently regulates oocyte function based on glucose conditions,these results suggest that oocytes originating from an obeseindividual may respond differently to energy substrates pre-sent in IVM medium. It may be important to optimize the invitro environment to better accommodate oocytes fromobese females, to best support oocyte competence andsubsequent developmental potential.

MATERIALS AND METHODS

Experimental Design

Effect of leptin on oocytes and cumulus cellsmatured in a physiological glucose environment Todetermine the effect of leptin on meiotic maturation andembryonic development of oocytes matured in a physiolo-gical glucose environment (2mM, G2; follicular fluid glu-cose concentration in normal weight pigs is �2mM; Bradet al., 2003), concentrations of 0, 10, and 100ng/ml (2G0L,2G10L, and 2G100L treatment groups, respectively) wer-esupplemented during IVM (physiological serumleptin concentration in pigs ranges between 5 ng/ml inlean sows to 20 ng/ml in obese sows; R.L.K., unpublisheddata). Next, the effect of leptin on oocyte and cumulus cellexpression of IRS1, IGF1, PPARg, and IL6 was examined.GLUT1 expression was determined in cumulus cells only.

Effect of leptin on oocytes and cumulus cellsmatured in a supraphysiological glucose environ-ment To determine the effect of leptin on meiotic matura-tion and gene expression of oocytes and cumulus cellsmatured in a supraphysiological glucose environment(50mM, G50), leptin was supplemented to maturationmedium at concentrations of 0, 10, and 100 ng/mL(50G0L, 50G10L, and 50G100L, respectively). The samegenes were analyzed as described above. Oocytesmatured in 5mM glucose without leptin (5G0L) wereincluded as a control to evaluate any changes due to theincrease in glucose concentration alone. This glucose con-centration is a 10-fold reduction in the supraphysiologicalconcentration, and is also the concentration typically usedin porcine IVM media (Funahashi et al., 2008).

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Effect of leptin during in vitromaturation on oocyteglucosemetabolism To determine the effect of leptin onoocyte glucosemetabolism through glycolysis and the PPPin conditions of standard (5mM) and supraphysiologicalglucose (50mM) concentrations, leptin was supplementedduring IVM at the same concentrations used previously (0,10, and 100ng/ml), and mature oocytes were selected formetabolic analysis immediately following IVM. For oocytesmatured in supraphysiological glucose, oocytes matured instandard glucosewithout leptin (5G0L) were again includedto determine alterations in oocyte metabolism due to theincrease in glucose concentration alone.

Oocyte Collection and In Vitro MaturationUnless specified otherwise, all chemicals were from

Sigma–Aldrich (St Louis, MO). Sow ovaries were collectedat a local abattoir. Follicles (3–9mm)were aspiratedwith an18-gauge needle attached to a vacuum pump. Oocytessurrounded by a minimum of two layers of unexpandedcumulus cells with a uniform cytoplasm were selected forIVM.COCswere thenwashed inHEPES-buffered syntheticoviductal fluid, SOF-HEPES (Gandhi et al., 2000), andcultured (7% CO2 in air) for 42 hr at 38.7�C in 500mlmaturation medium in a four-well plate (25–50COCs/well; Nunclon, Nalge Nunc Intl, Rochester, NY). Maturationmedium was Purdue Porcine Medium-modified for oocytematuration (PPMmat; Herrick et al., 2003), containing100 ng/ml epidermal growth factor, 0.01U/ml each porcineluteinizing hormone, and follicle-stimulating hormone(Sioux Biochemical, Sioux Center, IA), ITS (0.5mg/ml insu-lin, 0.275mg/ml transferrin, 0.25 ng/ml sodium selenite),0.5mg/ml recombinant human albumin (G-MM; Vitrolife,Kungsbacka, Sweden), and 0.2% fetuin.

Determination of Meiotic StageAftermaturation, oocytesweredenudedof cumulus cells

by vortexing in SOF-HEPES with 80–160U/ml hyaluroni-dase for 2.5min. Groups of approximately 20 oocyteswere mounted on glass slides, fixed in chloroform:aceticacid:ethanol (1:3:6) for 48 hr, stained with 1% (w/v) orceinin acetic acid, and examined for meiotic stage at 400�magnification. Oocytes that had progressed to the telo-phase or metaphase II stage were considered mature.

In Vitro Fertilization and Embryo CultureFollowing maturation, cumulus cells were removed as

described previously and oocytes were washed three timesin modified Tris-buffered medium (mTBM; Abeydeera andDay, 1997), supplemented with 2mM caffeine, 0.2% (w/v)fraction VBSA, and 1�PSA (100 IU/ml penicillin, 100mg/mlstreptomycin, 0.25mg/ml amphotericin B; MP Biomedicals,Solon, OH). Oocytes were placed into 50ml drops of mTBMunder 10ml mineral oil (20 oocytes/drop; Falcon 1007;Becton Dickinson Labware, Franklin Lakes, NJ). Spermwas prepared by placing 1ml of extended semen (1:5dilution, Androhep EnduraGuard, Minitube of AmericaInc., Verona, WI), warmed for 20min, onto a gradient

of 45%:90% Percoll� (GE Healthcare Life Sciences,Uppsala, Sweden) and centrifuged for 20min at 700g.The supernatant was removed and the remainingsperm pellet washed in 5.0ml D-PBS (GIBCO Invitrogen,Carlsbad, CA) twice by centrifuging for 5min at 1,000g.Spermwere then diluted inmTBM,and added to drops (finalvolume 100ml) containing oocytes for a final sperm con-centration of 250,000 sperm/ml. Gametes were coincu-bated for 5 hr in 6% CO2 in humidified air. Followingcoincubation, zygotes were washed three times and cul-tured in 50ml drops of NCSU-23 medium (10 zygotes/drop)containing 0.4%w/v crystallized BSA (MP Biomedicals)under 10ml mineral oil in 6% CO2, 10% O2, balance N2

for 6 days, when embryonic cleavage and blastocyst devel-opment were determined. Blastocysts were stained with0.01mg/ml Hoechst 33342 (Invitrogen, Carlsbad, CA) todetermine total blastocyst cell number under fluorescenceat 400� magnification.

Quantitative RT-PCR

Oocytes Transcript abundance of IRS1, IGF1, IL6, andPPARg were analyzed by quantitative real-time PCR(qPCR). The genes analyzed in this study were selectedbased upon their association with ovarian glucose meta-bolism and/or leptin function: IRS1, IGF1, and PPARg areinvolved in the insulin signaling pathway (Poretsky et al.,1999; Minge et al., 2008), GLUT1 is involved in glucoseuptake by the cumulus cells (Sutton-McDowall et al., 2010),and IL6 and PPARg are associated with leptin signaling innonreproductive tissues (De Vos et al., 1996; Walleniuset al., 2002). After the maturation period, oocytes weredenuded and the polar body visualized for determinationofmature oocytes.Oocyteswith apolar bodywere collectedfor analysis in three groups of 30 (Experiment 1) or 20(Experiment 2) oocytes. Total RNA was extract frompooled oocytes using PicoPure� RNA Isolation kit(Arcturus Biosciences, Mountain View, CA) according tothe manufacturer’s protocol. Residual genomic DNA waseliminated by treatment with RNase-Free DNase Set(Qiagen Inc., Valencia, CA). cDNA was generated usingSensiscript RT kit (Qiagen Inc.) according to themanufacturer’s protocol. Primers were designed as pre-viously described (Paczkowski et al., 2011). Primersequences of target and reference genes are presentedin Table 5. qPCR was performed as previously described(Fleming-Waddell et al., 2007; Paczkowski et al., 2011) on10-fold diluted sample cDNA run in duplicate. Target geneswere analyzed using iQ SYBR Green Supermix reagents(Bio-Rad, Hercules, CA).

For relative quantification, the threshold for eachtarget gene was adjusted to the threshold level ofthe reference gene glyceraldehyde 3-phosphate dehydro-genase (GAPDH, physiological glucose experiment) andhypoxanthine-guanine phosphoribosyltransferase (HPRT,supraphysiological glucose experiment), respectively.HPRT was used in the supraphysiological glucose experi-ment because GAPDH expression tended to be downre-

302 Mol Reprod Dev 79:296–307 (2012)

Molecular Reproduction & Development SILVA ET AL.

gulated (P¼0.10) with the increase in glucose from 5 to50mM. Reference genes were stably expressed (not sta-tistically different) between treatments within each experi-ment. Data were analyzed using the relative expressionsoftware tool, REST 2009 (Qiagen Inc.), as previouslydescribed (Paczkowski et al., 2011). Expression ratioswere generated using PCR efficiencies (E) of the targetand reference genes and the DCt values of the control andtreatment samples (Pfaffl, 2001). The threshold cycle (Ct)values represent the PCR cycle when the SYBR Greenfluorescence rises above the background fluorescence orthreshold. The levels of significance were calculated bypair-wise, fixed reallocation randomization tests and sig-nificance was determined with a P-value <0.05.

Cumulus cells Transcript abundance of IRS1, IGF1, IL6,PPARg, and GLUT1 were analyzed for cumulus cells col-lected from all oocytes present in a treatment per replicateafter denuding the oocytes, independent of maturationstate. Cumulus cells were removed to a 1.5-ml tube, thencentrifuged for 1min at 700g. After centrifugation, super-natant was removed and cumulus cells samples werefrozen at�80�C for qPCR analysis. Total RNA was extractfrom cumulus cells using PicoPure� RNA Isolation kit(Arcturus Biosciences, Mountain View, CA) according tothe manufacturer’s protocol. Residual genomic DNA waseliminated by treatment with RNase-Free DNase Set(Qiagen Inc.). The amount of RNA was quantified usingthe Quant-iT� RiboGreen kit (Molecular Probe, Eugene,OR) and 0.2mg was used for further reverse transcriptasereactions. For generation of cDNA, SuperScript� IIIReverse Transcriptase (Invitrogen Corporation, Carlsbad,CA) was used following the manufacturer’s protocol. Pri-mers design and qPCR were performed as describedabove.

Oocyte MetabolismAfter 42 hr of IVM, cumulus cells were removed, and

polar bodies were visualized to determine oocyte maturityas described; only mature oocytes were used in metabolicanalyses. Glycolysis and PPP activity were simultaneously

measured using [5-3H]glucose (0.013mM, 0.25mCi/ml;Perkin–Elmer NEN Life Sciences Inc., Boston, MA) and[1-14C]glucose (0.482mM, 0.0265mCi/ml; American Radi-olabeled Chemicals Inc., St Louis, MO), respectively(O’Fallon andWright, 1986; Rieger and Guay, 1988). Radi-olabeled glucose was dissolved in PPMmat-based meta-bolism medium containing 0.5mM unlabeled glucose forthe standard glucose experiment and 1.5mM unlabeledglucose for the supraphysiological glucose experiment. Inaddition, metabolism medium contained 1.5mM lactate,0.01mM pyruvate, and 1% recombinant human albumin(G-MM� Recombumin�, Vitrolife, Kungsbacka, Sweden).Oocytes were collected in 2ml metabolism medium, com-bined with 2ml metabolismmedium containing radiolabeledglucose, and placed in the cap of a 1.5-ml microcentrifugetube in a single 4ml drop. The cap containing the oocytewasthen replaced onto a 1.5-ml microcentrifuge tube filled with1.5ml 25mMNaHCO3, and the remaining air space gassedwith 5% CO2 in air as the lid was secured. Drops notcontaining an oocyte (blanks) were included in each assaytomonitor nonspecific releaseof 14Cand 3H. In addition, thetotal amount of radioactivity per tube was determined ineach assay by thoroughly mixingmetabolism drops withoutoocytes with the NaHCO3 trap. After a 3 hr incubation(37�C), the caps were removed, and 1ml of the NaHCO3

trap was added to a scintillation vial containing 200ml 0.1MNaOH, and the vials stored for 24 hr at 4�C. Tenmilliliters ofscintillation fluid (Ecolite�; MP Biomedicals) was added toeach vial, vials were shaken briefly, and then held at roomtemperature for at least 24 hr beforemeasuring radioactivityon a liquid scintillation counter.When counts for one or bothof the pathwayswere less than themean value of the blanksfor either label, the oocyte was considered non-viable andnot included in the analysis.

Statistical AnalysisFor nuclear maturation, each oocyte was assigned as 1,

if it had achieved the desired stage ofmaturation (telophaseor metaphase II), or 0 if it had not. For embryonic cleavageand blastocyst development data, the total numberof embryos that cleaved or developed to the blastocyst

TABLE 5. Details of the Primers Used for Quantitative Real-Time PCR Analysis

Gene Primer sequences (50–30) Accession number Amplicon size (bp)

GAPDHForward: ACATCAAGAAGGTGGTGAAG

AF017079 151Reverse: ATTGTCGTACCAGGAAATGAG

HPRTForward: GTGATAGATCCATTCCTATGACTGTAGA

AF143818 104Reverse: TGAGAGATCATCTCCACCAATTACTT

IGF1Forward: CTGGACCTGAGACCCTCTGT

NM_214256 150Reverse: GGAAGCAGCACTCATCCAC

IRS1Forward: GTTTCGTGAAGCTGAACTCG

EU681268 169Reverse: ATATTCTGGGCCACCACAG

IL6Forward: CGCAGCCTTGAGGATTTC

NM_214399 121Reverse: CCCAGTGGACAGGTTTCTG

PPARgForward: ATCAAGCCCTTCACCACTGT

NM_214379 153Reverse: GGACACAGGCTCCACTTTG

GLUT1Forward: ACTCCACAAGCATCTTCGAG

EU012358 150Reverse: CAGCCAGGCCTATGAGGT

Mol Reprod Dev 79:296–307 (2012) 303

LEPTIN’S EFFECT IN OOCYTES IS GLUCOSE DEPENDENT

stage in each drop was recorded (n¼ 12–14 drops,�10 presumptive zygotes/drop). Maturation, embryoniccleavage and blastocyst development were analyzed usingthe generalized linear mixed model (GLIMMIX) macro inSAS (SAS Institute Inc., Cary, NC) as previously described(Herrick et al., 2006); treatment was included in themodel as a fixed factor, and replicate as a random factor.For the continuous data (glucose metabolism and blasto-cyst cell number), the PROC MIXED procedure in SASwas used as previously described (Larson et al., 2011).Data were normalized using square root transformation.‘‘Treatment’’ was considered a fixed factor, and ‘‘replicate’’and ‘‘treat� replicate’’ were considered random factors.When the explanatory variable was significant, Tukey–Kramer’s test was used for multiple comparisons betweentreatments. Differences were considered significant atP�0.05, and differences with 0.05<P�0.10 were con-sidered statistical tendencies.

ACKNOWLEDGMENTS

The authors would like to acknowledge support from theUniversity of Illinois, College of Agriculture, Consumer andEnvironmental Sciences. We acknowledge the assistanceof Dr. Jason Herrick for his help with editing this article.

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LEPTIN’S EFFECT IN OOCYTES IS GLUCOSE DEPENDENT